Environmental Contamination of Chrysotile Asbestos ... - Springer Link

Arch. Environ. Contam. Toxicol. 47, 281–289 (2004) DOI: 10.1007/s00244-004-3161-7

A R C H I V E S O F

Environmental Contamination a n d Toxicology © 2004 Springer ScienceⴙBusiness Media, Inc.

Environmental Contamination of Chrysotile Asbestos and Its Toxic Effects on Growth and Physiological and Biochemical Parameters of Lemna gibba A. K. Trivedi,1 I. Ahmad,1 M. S. Musthapa,1 F. A. Ansari,1 Q. Rahman1 1

Fibre Toxicology Division, Industrial Toxicology Research Centre, P. B. No. 80, M. G. Marg, Lucknow 226 001, India

Received: 23 July 2003 /Accepted: 23 February 2004

Abstract. Asbestos was monitored in water, sediment, and aquatic plant samples around an asbestos cement factory. Based on asbestos concentration found in aquatic plants during monitoring, and the propensity of asbestos to cause oxidative stress in animal models, laboratory experiments were conducted to assess toxicity of chrysotile asbestos on an aquatic macrophyte, duckweed (Lemna gibba). L. gibba plants were exposed to two concentrations of chrysotile asbestos (0.5 ␮g and 5.0 ␮g chrysotile in 5.0 ␮l double distilled water) twice per week during a period of 28 days and cultured in medium containing 0.1 g chrysotile/L. Control plants were cultured in medium without chrysotile asbestos. Effect of chrysotile exposure on certain growth and physiological and biochemical parameters was evaluated. An inhibition effect of chrysotile exposure was found on the number of fronds, root length, and biomass. Similar alterations in contents of chlorophyll, carotenoid, total free sugar, starch, and protein were also found. Contrary to effect on these parameters, a dose- and timedependent increase in efflux of electrolytes, lipid peroxidation, cellular hydrogen peroxide, catalase, and superoxide dismutase activity was found. The results indicate oxidative stress and phytotoxicity of chrysotile asbestos on duckweed.

Asbestos fibers are divided into two classes, chrysotile and amphibole, on the basis of their crystal structure (Light and Wei 1977). Crysotile is a fibrous hydrated magnesium silicate mineral [Mg3Si2O5(OH)4] that is used in approximately 3000 commercial products (Ramanathan and Subramanian 2001). Although its use is now banned in some developed countries, it is still in use in developing countries. In India, several states have many asbestos industries, 60% of which are in operation, and production is approximately 2000 tons/mo (Ramanathan and Subramanian 2001). Air pollution levels of asbestos were reported to be increased in the areas surrounded by asbestos industries (International Programme on Chemical Safety [IPCS] 1998). Because of difficulties in decreasing the emission of fine particles of asbestos during factory operation, the

Correspondence to: Q. Rahman; email: qrahman_itrc.yahoo.ca.im

particles are released into the environment. Therefore, monitoring and analysis of biotic and abiotic samples in the nearby ecosystem can address many questions about source, distribution, partitioning, and transport of asbestos. Under natural conditions, chrysotile fibers can be transported by wind and water (IPCS 1998). Natural fresh water is an important receptor of many toxic substances released by industrial, agricultural, and domestic activities (Kumari et al. 2001). Various health effects in man and laboratory mammals have been documented due to chrysotile fiber exposure (Hauptman et al. 2002). However, the potential ecologic impacts of this material have largely been ignored (National Institute 1989). Furthermore, one of the great generalizations of cell biology is that the cells of higher organisms, whether from plants or animals, are fundamentally similar (Prescott 1982). Chrysotile fibers carry a positive surface charge at pH ⬍11.8 (Speil and Leinweber 1969). These charged fibers presumably would be attracted to negatively charged protein groups in cell membranes. The chrysotile fiber would then be surrounded by proteins and submerge into the cell. This series of events could be a mechanism by which fibers could gain entry into the cell (Harington et al. 1975). Moreover, a common effect of many pollutants is to generate free radicals and reduced forms of oxygen, which may damage cellular components such as lipids, proteins, or DNA (Foyer et al. 1997). The importance of reactive oxygen species (ROS) in contributing to asbestos cytotoxicity has been reported by several investigators (Kamp et al. 1992). Several studies have shown that fiber– cell interaction is not necessary for the production of ROS; for example, chrysotile and crocidolite asbestos in cell-free solutions of water or physiologic saline spontaneously generate superoxide (O•2) or hydroxyl radical (OH•⫺) (Ebenhardt et al. 1985). Duckweed is an important food species for aquatic herbivores; is a good dietary supplement and nutrient source (Oron et al. 1985) for humans (Majid et al. 1984), livestock, and fish (Lehman et al. 1981); and is used as a good fertilizer supplement (Mabagwu and Adeniji 1988) and indicator of water pollution (Nasu and Kugimoto 1981). It is widely recommended for aquatic toxicity studies (Environmental Protection Agency [EPA] 1985). The selection of exposure concentration is based on the range of fibers found during monitoring (Rahman et al. 2001). In this study, we investigated the effect of

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chrysotile asbestos in the vicinity of an asbestos cement factory and its toxic effects on growth as well as physiological and biochemical parameters of a primary producer, L. gibba.

Materials and Methods Asbestos Fiber Analysis Asbestos analysis in different samples was carried out following the method of American Public Health Association (APHA et al. 1998), United States Environmental Protection Agency (USEPA 1993), and Indian Standards [IS] (1986). Water, sediment, and plant samples were collected from different locations near an asbestos cement factory located at Mohanlalganj, which is approximately 25 km from Lucknow, Uttar Pradesh (India). Sediment and plant samples were dried at 65°C until constant weight was obtained, and further processing was done in dried material. Sediment and plant samples were ashed separately according to site of collection at 500°C for 2 hours in muffle microwave, and ash was mixed with nitric acid and further diluted with deionized water to avoid any damage to the Millipore filter during filtration process. The pond water samples, processed sediment, and plant samples were filtered through a Millipore membrane filter paper with a pore diameter of 0.8 ␮m (catalogue no. AABP 04700; Millipore Corporation, Bedford, MA), which retains asbestos fibers present in the samples. The fibers are subsequently transferred on a slide and made transparent by the addition of 200 to 300 ␮l standard immersion oil (Carl Zeiss, Oberkochen, Germany). Transparent slides were air dried and used for asbestos analysis by phase-contrast polarized microscopic method (IS 1986). Length of asbestos fibers was measured in the range of ⬍10, 11 to 20, and ⬎20 ␮m, and a relative count of fibers was also estimated in the original material.

A. K. Trivedi et al.

Physiologic Parameters Fresh tissues were extracted in 80.0% acetone for the spectroscopic estimation of chlorophyll (Strain et al. 1971) and carotenoid (Duxbury and Yentshe 1956) content. Total free sugar was estimated colorimetrically in alcoholic extract (Montgomery 1957). Residue left after alcoholic extraction was hydrolyzed with perchloric acid (Agrawal et al. 1977), and starch was estimated as free sugar. The total protein content in fresh fronds was estimated by folin ciocalteau reagent method (Lowry et al. 1951). Electrolyte efflux was measured per the procedure described by Dionisio-Sese and Tobita (1998).

Biochemical Parameters To measure lipid peroxidation, the thiobarbituric acid test, which determines malondialdihyde (MDA) as an end product of lipid peroxidation (Heath and Packer 1968), was used per the procedure described by Dhindsa and Matowe (1981). The cellular hydrogen peroxide content was measured by fluorometrical assay with hemovanillic acid according to Ishikawa et al. (1993; with some modifications). Catalase activity was determined by consumption of hydrogen peroxide (Rao et al. 1996). Superoxide dismutase activity, the basis of which is its ability to inhibit the photochemical reduction of nitroblue tetrazolium (Beauchamp and Fridovich 1971), was assayed per the procedure described by Stewart and Bewley (1980).

Statistical Analysis Data for each parameter were evaluated for statistical significance using two-way analysis of variance (ANOVA) to compare the means considering duration of exposure and concentration as independent variables. The individual treatment between the two groups was assessed by computation of least significant difference taking t values for error df at the 5% level of significance.

Collection and Culture of Lemna gibba To study toxic effects of chrysotile asbestos on Lemna gibba, plants were collected from the natural habitat in an aquatic body, washed axenically, and maintained in Hoagland medium (EPA 1975) in the laboratory under a light-to-dark period of 16 h:8 h and controlled humidity (60%). The young plants of the third generation were transferred to sterilized petri dishes and used for experiment. Chrysotile fibers ⬍30 ␮m were used in the study. Two suspensions of chrysotile fibers were prepared by adding 0.1 g and 1.0 g chrysotile separately in 1.0 L double-distilled water while stirring constantly. Five ␮l suspension was applied per frond twice per week. The plants were cultured in a medium containing 0.1 g chrysotile/L to reproduce conditions similar to the field, where chrysotile enters in the aquatic system with drainage water as well as falls through on fronds. Control plants were cultured in nutrient medium without chrysotile fiber. Experiments were conducted in 5 replications. Five petri dishes having 20 plants in each were maintained per replicate. Data presented are mean of 3 independent experiments.

Growth Parameters Effect on increase in total number of fronds, total fresh weight of plants (biomass) per petri dish, and root length per plant was evaluated at different time intervals after exposure.

Results and Discussion Asbestos contamination was found in all of the sediment samples collected and analyzed, but more fibers were present in samples collected from the west and south corners of the pond (Table 1). This may have occurred because of the vicinity of these corners to the factory. Furthermore, the lengthwise distribution of fibers was quite varied; more large fibers were found in pond areas close to the factory compared with smaller fibers because smaller fibers can travel greater distances in air and water. These data further explain the fact that the large fibers are removed from air and water by gravitational settling at a rate depending on their size, but smaller fibers may remain suspended for long periods of time. Similarly, in plant samples more small fibers were present, and fiber count per gram dry weight was more in root compared with other plant parts (Table 2). This may have occurred because of continuous contact of roots with contaminated sediment or water. According to Toxic Release Inventory ’99 the total release of asbestos into the environment (including air, water, and soil) from 87 facilities was 13.6 million pounds (TRI 99 2001). Asbestos fibers do not endure significant transformation and degradation in soil (Musthapa et al. 2003), which makes it easy to move from one trophic level to another in the ecosystem. This situation high-

Chrysotile Asbestos and Lemna gibba

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Table 1. Asbestos residue in pond water and sediment samples near an asbestos cement factory

Sample Pond Pond Pond Pond Pond Pond Pond Pond a

water east water west water north water south sediment east sediment west sediment north sediment south

% Fiber (Lengthwise)

Total fiber/L (or g dw)

⬍10 ␮m a

287 298 282 304 399 404 360 420

19.86 (57) 11.07 (33) 7.80 (22) 11.84 (36) 3.01 (12) 4.95 (20) 5.00 (18) 4.52 (19)

11–20 ␮m

⬎20 ␮m

16.37 (47) 17.78 (53) 15.60 (44) 12.89 (39) 11.02 (44) 10.40 (42) 13.61 (49) 9.29 (39)

63.76 (183) 71.14 (212) 76.59 (216) 75.33 (229) 85.96 (343) 84.65 (342) 81.38 (293) 86.19 (362)

Figures in parentheses indicate number of fibers out of total number.

Table 2. Asbestos burden in aquatic plants and their parts around asbestos cement factory % Fiber (Lengthwise) Name of Plant Nelumbo nucifera

Nymphaea nouchali

Ranunculus scleratus

Lemna gibba a

Name of Part Root Pedicel Leaves Root Pedicel Leaves Root Stem Leaves Total

Total Fiber/g dw 41 34 24 47 42 38 44 25 23 21

⬍10 ␮m a

40.78 (20) 50.00 (17) 50.00 (12) 48.93 (23) 47.61 (20) 65.78 (25) 40.90 (18) 48.00 (12) 52.17 (12) 57.14 (12)

11–20 ␮m

⬎20 ␮m

29.26 (12) 29.41 (10) 33.33 (8) 29.78 (14) 30.95 (13) 26.31 (10) 34.09 (15) 32.00 (8) 34.78 (8) 28.57 (6)

21.95 (9) 20.58 (7) 16.66 (4) 21.27 (10) 21.42 (9) 7.89 (3) 25.00 (11) 20.00 (5) 13.04 (3) 14.28 (3)

Figures in parentheses indicate number of fibers out of total number.

lights the need to study the transformation, persistence, and effect of asbestos on different growth and metabolic processes of primary producers, such as L. gibba, because it has a doubling time of only 0.7 day (Hughes et al. 1988) and is widely recommended for aquatic toxicity testing (USEPA 1985). The reason behind conducting 28 days of experiments was to assess the long-term effects of environmental contamination of asbestos. Figure 1 illustrates the inhibition effect of chrysotile asbestos on number of fronds on postexposure day 3. A decrease in number of fronds per petri dish (21.78% and 25.75%, respectively) was found in plants exposed to 0.5 and 5.0 ␮g chrysotile/frond. These figures gradually increased to significant values of 39.82% and 49.34%, respectively, on postexposure day 28. Root length was found to be less sensitive to chrysotile exposure; effect on root length was significant only on postexposure days 21 and 28. Initially on postexposure day 3, 10.66% and 13.78% reductions in root length at 0.5 and 5.0 ␮g chrysotile/frond, respectively, were found. On postexposure day 28, a decrease in root length (36.41% and 46.58%, respectively) was found in fronds exposed to 0.5 and 5.0 ␮g chrysotile (Figure 2). A significant decrease in biomass in fronds exposed to 0.5 and 5.0 ␮g chrysotile/frond found after an exposure period of 14 days and 7 days, respectively. At postexposure day 28, the decrease in biomass was 32.68% and 44.08%, respectively, in plants exposed to 0.5 and 5.0 ␮g chrysotile/frond (Figure 3). Chrysotile is stable in alkaline water, but magnesium leach-

ing occurs from fiber structure under acidic conditions. Chrysotile’s surface charge changes from positive in alkaline conditions to negative under acidic conditions (due to loss of magnesium ion from surface brucite layers) (IPCS 1998). It is unstable in water, physiologic saline, and acidic medium because it loses its magnesium ion from the fiber structure (Hodgson and Jones 1986). Leaching of magnesium and trace metals present in fiber composition might locally increase levels of these elements (Schreier and Timmenga 1986). The inhibition effect on number of fronds, root length, and biomass (i.e., growth inhibition) might be due to the toxicity of magnesium and other trace metals leached from chrysotile fibers because the magnesium content of chrysotile is related to its toxicity (Light and Wei 1977). Moreover, fiber size and geometry appear to be the main issue for human health, but bulk and trace metal chemistry have been identified as factors and agents detrimental to plant growth (Roberts and Proctor 1993). Chrysotile exposure caused a significant decrease in chlorophyll content during the entire period of 28 days. At postexposure day 3, 2.66% and 7.89% decreases in the chlorophyll content of fronds exposed to 0.5 and 5.0 ␮g chrysotile/frond, respectively, were found, and this increased to 19.08% and 31.31%, respectively, on postexposure day 28 (Figure 4). Although the decrease in carotenoid content was less, i.e., 18.65% and 29.31%, on postexposure day 28 in fronds exposed to 0.5 and 5.0 ␮g chrysotile/frond, it was in conformity with the effect on chlorophyll content (Figure 5). Chlorophyll and carotenoid are plant pigments that play a crucial role in the

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Fig. 1. Effect of chrysotile on number of fronds in duckweed. Values represent means ⫾ SD expressed as percent controls. Control values were 1.0, 8.20, 12.60, 18.20, and 24.00 on postexposure days 3, 7, 14, 21, and 28, respectively. *Denotes statistically significant difference from respective control values (p ⬍ 0.05)

Fig. 2. Effect of chrysotile on root length in duckweed. Values represent means ⫾ SD expressed as percent of respective controls. Control values were 2.60, 6.20, 7.00, 7.60, and 8.60 cm on postexposure days 3, 7, 14, 21, and 28, respectively. *Denotes statistically significant difference from respective control values (p ⬍ 0.05)

Fig. 3. Effect of chrysotile on biomass in duckweed. Values represent means ⫾ SD expressed as percent of respective controls. Control values were 6.64, 23.33, 40.33, 58.30, and 74.26 mg on postexposure days 3, 7, 14, 21, and 28, respectively. *Denotes statistically significant difference from control values (p ⬍ 0.05)

photosynthetic conversion of radiant energy into chemical energy. A significant decrease in these pigments may be related to the effect of chrysotile on macromolecules and enzymes because several biological macromolecules and enzymes are known to be adsorbed on the surface of asbestos and thus affect the physiologic function of cells (Fisher et al. 1987). In addition, suspended chrysotile fibers may adsorb organic materials, which eventually cover the entire fiber surface (Bales and Morgan 1985) and affect cell function. The carotenoids, particularly ␤-carotene, are isoprene-based pigments. In addition to acting as accessory pigments in photosynthesis, carotenoids also function as scavengers of 1O2 and quenchers of tripletstate chlorophyll molecules (Young 1991). A decrease in carotenoid content showed decreased ability of plants to cope with oxidative stress (Farooq et al. 2000). Contrary to the effects on the previously mentioned parameters, efflux of electrolytes from roots was also increased by

22.19% and 35.05% on postexposure day 3 and gradually increased to 203.97% and 262.11% on postexposure day 28 in plants exposed to 0.5 and 5.0 ␮g chrysotile/frond, respectively (Figure 6). This indicates massive damage to cellular membranes and loss of cellular constituents (Dionisio-Sese and Tobita 1998). Furthermore, a significant decrease in protein, starch, and total free-sugar content was found (Table 3). The decrease in protein content varied from 13.48% and 23.39% on postexposure day 3 to 38.94% and 50.50% on postexposure day 28 in 0.5 and 5.0 ␮g/frond chrysotile– exposed plants, respectively. A similar decrease in total free-sugar content was also found: 6.73% and 30.73% on postexposure day 3, increasing to 26.57% and 44.92% on postexposure day 28 in 0.5 and 5.0 ␮g/frond chrysotile– exposed plants, respectively. The decrease in starch content was 6.90% and 19.44% on postexposure day 3, which increased to 22.57% and 35.57% on post-


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Fig. 4. Effect of chrysotile on chlorophyll content in duckweed. Values represent means ⫾ SD expressed as percent of respective controls. Control values were 0.891, 1.020, 1.258, 1.380, and 1.385 mg/g fresh tissue on postexposure days 3, 7, 14, 21, and 28, respectively. *Denotes statistically significant difference from control values (p ⬍ 0.05)

Fig. 5. Effect of chrysotile on carotenoid content in duckweed. Values represent means ⫾ SD expressed as percent of controls. Control values were 0.258, 0.280, 0.311, 0.319, and 0.321 mg/g fresh tissue on postexposure days 3, 7, 14, 21, and 28, respectively. *Denotes statistically significant difference from control values (p ⬍ 0.05)

Fig. 6. Effect of leakage of electrolytes in duckweed. Values represent means ⫾ SD expressed as percent of controls. Control values were 21.33, 26.67, 29.67, 32.36, and 35.67 ␮mhos on postexposure days 3, 7, 14, 21, and 28, respectively. *Denotes statistically significant difference from control values (p ⬍ 0.05)

exposure day 28 in fronds exposed to 0.5 and 5.0 ␮g chrysotile/ frond, respectively. The significant decrease in protein, total free sugar, and starch content in chrysotile-exposed plants may be due to the explicitly unknown roles of bulk and trace metals involved in fiber structure and chemistry. A number of metals—including nickel, chromium, antimony, and cobalt—are associated with asbestos fibers, which have been implicated as one of the causative factors of asbestos-related diseases in animals (Barbeau et al. 1985) and may be toxic to plants also. It is currently assumed that the negative effect of the various environmental stresses are at least partially caused by the generation of ROS and/or the inhibition of the system that defends against them (Shalata and Tal 1998). Lipid peroxidation gradually increased from 1.43% and 9.91% on postexposure day 3 to 31.12% and 33.46% on postexposure day 28 in 0.5 and 5.0 ␮g/frond chrysotile– exposed plants, respectively, which indicates oxidative stress and membrane

damage (Figure 7). ROS brought about peroxidation of membrane lipids and led to membrane damage (Scandalios 1993). Because lipid peroxidation is the symptom most easily ascribed to oxidative damage (Zhang and Kirkham 1996), it is often used as an indicator of increased oxidative stress (Jagtap and Bhargava 1995). Another parameter of oxidative stress is hydrogen peroxide, which is formed during various oxidase reactions and can lead to highly reactive oxygen radicals (Halliwell and Gutterdge 1989). Hydrogen peroxide is involved in oxidative stress signaling (Morita et al. 1999). Cellular hydrogen peroxide content was increased by 3.61% and 35.06% on postexposure day 3 and 68.87% and 96.55% on postexposure day 28 in 0.5 and 5.0 ␮g/frond chrysotile-exposed plants, respectively (Figure 8). Previously, a similar increase in hydrogen peroxide content was reported in response to different environmental stresses such as chilling (Fadzillah et al. 1996), ultraviolet radiation

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Table 3. Effect of chrysotile exposure on protein, total free-sugar, and starch content in duckweed Postexposure Days Protein content (mg/g fresh wt) 3 7 14 21 28 Total free sugar (mg/g fresh wt) 3 7 14 21 28 Starch (mg/g fresh wt) 3 7 14 21 28

Control

0.5 ␮g/frond

0.5 ␮g/frond

39.76 ⫾ 2.098 39.66 ⫾ 2.173 40.52 ⫾ 1.639 39.56 ⫾ 1.417 39.40 ⫾ 1.528

34.40 ⫾ 1.258* 32.96 ⫾ 0.993* 30.32 ⫾ 0.825* 27.58 ⫾ 0.901* 24.06 ⫾ 0.939*

30.46 ⫾ 0.492* 28.10 ⫾ 0.696* 25.20 ⫾ 0.815* 22.24 ⫾ 0.853* 19.50 ⫾ 0.689*

0.6208 ⫾ 0.0085 0.6240 ⫾ 0.0114 0.6180 ⫾ 0.0057 0.6170 ⫾ 0.0044 0.6210 ⫾ 0.0065

0.5790 ⫾ 0.0089* 0.5580 ⫾ 0.0083* 0.5290 ⫾ 0.0061* 0.4970 ⫾ 0.0083* 0.4560 ⫾ 0.0238*

0.4300 ⫾ 0.0158* 0.4120 ⫾ 0.0083* 0.3900 ⫾ 0.0079* 0.3680 ⫾ 0.0057* 0.3420 ⫾ 0.0125*

70.46 ⫾ 0.6503 69.66 ⫾ 1.4553 70.30 ⫾ 1.2083 69.80 ⫾ 1.2165 69.64 ⫾ 0.7668

65.60 ⫾ 0.9165* 63.28 ⫾ 0.5495* 60.68 ⫾ 0.8871* 57.68 ⫾ 0.5805* 53.92 ⫾ 0.7463*

56.76 ⫾ 0.5029* 53.00 ⫾ 0.8831* 51.12 ⫾ 0.7987* 48.12 ⫾ 0.9038* 45.56 ⫾ 0.7635*

Note: Data represent means of three independent experiments. Values represent means ⫾ SD. * Denotes statistically significant difference from respective control values (p ⬍ 0.05). wt: weight.

Fig. 7. Effect of chrysotile on lipid peroxidation in duckweed. Values represent means ⫾ SD expressed as percent of control. Control value was 1.67 ⫾ 0.081 nmol MDA formed/mg protein/h. *Denotes statistically significant difference from control values (p ⬍ 0.05)

Fig. 8. Effect of chrysotile on hydrogen peroxide content in duckweed. Values represent means ⫾ SD expressed as percent of control. Control value was 2.33 ⫾ 0.067 nmol hydrogen peroxide formed/mg protein. *Denotes statistically significant difference from control values (p ⬍ 0.05)

(Murphy and Huerta 1990), heat (Dat et al. 1998), and excessive light (Karpinski et al. 1997). Catalase is thought to lower oxidative damage by converting hydrogen peroxide to water and oxygen (Scandalios et al. 1997). The present study reveals that catalase activity was increased by 1.64% and 6.15% on postexposure day 3 and 22.66% and 48.89% on postexposure day 28 in 0.5 and 5.0 ␮g/frond chrysotile–

exposed plants, respectively (Figure 9). Overexpression of the gene for this enzyme protects leaves against ROS (Zelitch et al. 1991), while catalase-deficient plants are more sensitive to various stresses (Willekens et al. 1997). In the present study, superoxide dismutase (SOD) activity increased by 3.61% and 20.31% on postexposure day 3 and 39.45% and 71.65% on postexposure day 28 in 0.5 and 5.0


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Fig. 9. Effect of chrysotile on catalase activity in duckweed. Values represent means ⫾ SD expressed as percent of control. Control value was 53.43 ⫾ 1.76 ␮mol hydrogen peroxide decomposed/min/mg protein. *Denotes statistically significant difference from control values (p ⬍ 0.05)

Fig. 10. Effect of chrysotile on superoxide dismutase activity in duckweed. Values represent means ⫾ SD expressed as percent of control. Control value was 7.89 ⫾ 1.09 enzyme U/mg protein. *Denotes statistically significant difference from control values (p ⬍ 0.05)

␮g/frond chrysotile– exposed plants, respectively (Figure 10). The scavenging of superoxide by SOD is an important mechanism to cope with stress condition and has been studied extensively (Bowler et al. 1992). Industrial production of asbestos products causes environmental contamination, and asbestos fibers reach to biota and abiota in the vicinity, a phenomenon also found in our earlier study (Musthapa et al. 2003). Chrysotile-exposed plants have shown stress symptoms in conformity with the previous report (IPCS 1998). In conclusion, chrysotile causes growth inhibition; decreased photosynthetic pigments, protein, total freesugar and starch content; and increased efflux of electrolytes, lipid peroxidation, cellular hydrogen peroxide levels, and catalase and SOD activity in chrysotile-exposed plants. Results indicate that chrysotile asbestos causes oxidative stress–mediated toxicity in plants.

Acknowledgments. Funding was provided by the Ministry of Environment and Forests (Government of India), New Delhi. The authors thank P.K. Seth for his keen interest in this study and M. Ashquin for skillful technical assistance.

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